Do a checking gel to see if you have an insert amplifying in each sample (if there is a ~200bp band, that means there is no insert; if no band, means there was no or very few cells in the boiled solution). Use 3µl of PCR product and 2µl of loading dye for the checking gel. If clean single band of the expected insert size, do the standard next steps for sequencing (Agencourt/Beckman prep, or if UCLA core – Exosap and Big Dye rxn).<br>

Lab safety

You must always wear closed-toes shoes and a lab coat when in the lab!

Always use gloves when working in the lab but DO NOT WEAR GLOVES OUTSIDE OF THE LAB (e.g. hallway, elevator, etc).

Pre-PCR is carried out in room 4146. All post-PCR activity takes place in room 4162. Discard gloves when moving between rooms and keep all equipment in their designated rooms. AVOID moving things back into the post-PCR room 4162. These precautions are to avoid contamination problems.

If you use the last of the supplies (such as tips or gloves), please restock these items.

When stock solutions, kits, gloves, Kimwipes, etc runs out, it is your responsibility to make sure these get replaced. If more needs to be ordered, please notify Sarah.

At the end of the day, remove all of your stuff from the bench you were using and return all equipment to their appropriate places, washing glassware with deionized water and hanging on the rack to dry.

Do not prop open the laboratory doors. Many of the researchers keep personal belong- ings in the office/lab. We want to prevent theft and follow Environmental Health and Safety guidelines.

Please turn off equipment if it is not in use. Especially hot plates! Make sure they are off when you are done using them.

If you uncomfortable at all with any laboratory technique, please do not attempt the procedure and ask for guidance.

Your supervisors (graduate student or post-doctoral researcher and Dr. Wayne) are responsible for you. Please make every effort to be a good laboratory citizen. The Wayne lab is filled with friendly and knowledgeable people. Please ask questions when you are at all uncertain about your technique, procedure, or project. Please make every effort to attend our weekly laboratory meetings, as this is an important forum for meeting the lab members and keeping up-to-date on issues in the lab (i.e. new techniques, current research projects underway, presenting your own work, etc.). This is also the time for the rest of the Wayne lab to meet you, so they are familiar with you and are able to provide help if needed.

Recipes for the general-use buffers

5M Sodium Chloride (NaCl) (makes 250mL)
1. Obtain a beaker with at least 250mL capacity
2. Measure out 73.05g of solid sodium chloride (NaCl) and add to beaker
3. Add distilled water up to total volume of slightly less than 250mL
4. Place beaker on stirring block, add stir bar
5. Stir and heat slightly
6. Add water until total volume is 250mL and sodium chloride is totally dissolved
7. Remove stir bar with magnet and place solution in a 500mL bottle with a loosely fitted cap
8. Autoclave (liquid cycle: ~20 mins)
9. Once cooled, run solution through sterile filter system

Obtain a short length of plastic hose to connect vacuum line to filter unit

Make sure receiving unit (bottom bottle) and filter unit (top half) are held together tightly

Qubit quantification of double-stranded DNA

Qubit gives a more accurate reading of the amount of dsDNA in a sample. Because we are using a High Sensitivity (HS) kit, your samples should be 1-100 ng/uL to ensure accuracy. You can use anywhere from 1-10uL of sample in each tube, always with a total volume of 200μL. We use 2μL to save sample and avoid potential pipetting errors of using 1μL.

The dye used in Qubit is light sensitive. Keep dye tube covered in aluminum foil. Put sample tubes with dye in tube rack covered with aluminum foil or in a draw when possible.

Materials

Qubit dsDNA HS Assay Kit

HS buffer

HS standard #2 (10ng/μL or 100ng/μL)

Quant-it dsDNA reagent (light sensitive dye)

Qubit Assay tubes

Tinfoil

Methods
1. Thaw High Sensitivity buffer, light-sensitive dye, standard and samples to room temperature (or use room temperature stock).
2. Cover a tube rack in aluminum foil for storing samples (or place in a drawer).
3. Pull Qubit tubes for number of samples plus two for the standards.
4. Label tubes STD-1 and STD-2, and label one tube per sample you will quantify. Put in tube rack covered in aluminum foil (or drawer).
5. In a sterile 1.5 mL tube covered in tinfoil mix the following. You should master mix this for the number of samples plus three to include standards and extra.

Nanodrop 2000 quantification of DNA

Please use your own materials (pipette, pipette tips, Kimwipes, etc). Start NanoDrop 2000 software and select analysis method (e.g. Nucleic Acid for DNA). Carefully apply a droplet of water (1-2µL) to the pedestal to clean and initialize the instrument. Use a Kimwipe to clean the pedestal and the top in between readings. At the prompt, name the results file and save it in the “My Documents” folder. Apply 1-2µL to the pedestal and press the “Blank” button (AE buffer, TE buffer, water, etc). Clean the pedestal and apply 1-2µL of sample, then press the “Measure” button. After measuring, the concentration and other data appear in the software window. Continue measuring remaining samples. After measuring, view the results table.

Polymerase Chain Reaction (PCR): General Introduction

A PCR contains the following necessary reagents:

PCR-buffer. Salt and pH-stabiliser. User stock of 10x is kept in your box.

MgCl2. Salt which is required for the Taq polymerase to work. The standard rxn concentration is 1.5 mM (range 1-4). Higher concentrations makes the Taq polymerase less specific and favours amplifications of short fragments. Too much MgCl2 often results in multiple bands. User stock of 25mM is kept in you.

dNTPs. Free nucleotides (Gs, As, Ts and Cs) of which the artificial DNA copies are made. User stock (10 mM of dNTP Mix, which has 2.5mM of each dNTP) is kept in your box.

Primers. Single stranded DNA (oligonucleotides), usually the length of 18-30 bp. Primers used for RAPD are normally shorter, 10-15 bp. Stock solutions at 100µM are normally, and user stocks at 10µM are stored in your PCR box.

Taq DNA polymerase. The enzyme that puts the free nucleotides together. It starts at the 3' end of the primer and uses the complementary DNA strand as a template. User stock of 5 units/µl is kept in your box.

Template DNA. The source of DNA for the PCR amplification. This could be DNA extracted from blood, skin, feathers, or old PCR products. We use a standard concentration at 25 ng/µl but depending on the organism and protocol, the concentration might need further adjustments (5-100 ng/µl).

In addition to these reagents, you may find that researchers are adding other reagents to their reactions in hope of getting better results. You may consider:

To set up a PCR:
1. Make sure you have reserved a thermocycler for your usage. The sign-up dry-erase board is in 5202.
2. Create a PCR protocol and calculate how much you need of your master mix/ cocktail (number of rxn + 10%). Don't forget the rxn for the blank (negative control). The master mix should contain everything except the template (but you should add the Taq just before aliquoting).
3. Make sure the PCR bench is clean.
4. Thaw template DNA and all reaction reagents on the PCR bench, except the Taq polymerase which should remain in the freezer until needed (it contains glycerol so it does not need thawing).
5. Program the PCR machine while the reagents are thawing.
6. Mix and spin all reagents and keep on ice (once experienced with the technique, keeping all reagents on ice is not necessary, unless there are unexpected delays).
7. Place an appropriate number of PCR tubes in a tray, and briefly label each row of tubes.
8. Aliquot your desired amount of template DNA prior to adding Taq polymerase to your cocktail.
9. Make the cocktail in an 1.5 ml eppendorff tube (snap cap). Start with adding the ddH20 and save the Taq DNA polymerase to the last step. Mix by using the 200 µl pipette set at 150µl and pipette up and down a few times.
10. Dispense an appropriate volume of master mix into each of the reaction tubes (total rxn minus amount of template DNA).
11. Add the template (change tip between samples!!!!).
12. Fix the lids on the tubes.
13. Bring the tubes over to the thermocyclers in room 4162 (on ice if you haven't programmed them yet--it may take a few minutes for a newly turned on machine to warm up). Start the PCR machine. Select and run your program, and ALWAYS USE A HEATED LID! Load the tubes, close and tighten the lid, then you are ready to go!

Programming the thermocycler (choosing a temperature profile):
The standard PCR starts with a warming up phase of 3 minutes at 94˚C, to make the template DNA single stranded (denatured). Then follows the cyclic phase that characteristically consists of three different steps.
1. 94˚C. This is again the denaturing step that initiates all cycles and is normally set between 30-60 sec.
2. 37-70˚C. The annealing temperature when the primer is allowed to settle on the template DNA. This step is usually set between 30-120sec. The chosen temperature depends on the melting temperature, Tm, of the primer (length and GC-content).
3. 72˚C. The elongation temperature is the optimal working temperature of the Taq DNA polymerase. This step is set between 5-500 sec depending on the length of the desired fragment. A rule of thumb is that the Taq polymerase builds about 1,000 nucleotides per minute.
4. The number of cycles used varies normally between 20-40 depending on the template DNA concentration, quality, length of product, and above all, empirical experience with the focal reaction.
5. The reaction is normally ended by a 10 minute phase at 72˚C. This will allow the Taq polymerase to add a protruding A at the 3' end of the fragments. This step is very important when cloning the PCR fragments by means of TA-cloning.

Recipe for 2% agarose gels:
1. If a 2% solution is already made up, proceed to step 3 to melt the gel. To make 300 ml of new solution, take a weighing boat from the drawer, place it on the balance, tare, and add 6.0 g agarose (in the cupboard of reagents behind the door). Add to flask.
2. Add 300 ml of 1x TBE buffer (in carboy).
3. Place the bottle in the microwave and boil for repeated 30 sec intervals until the agarose has melted (make sure it doesn't boil over!). Use the orange heat-protection gloves when handling the warm bottle.
4. Tape the ends of the gel casts and insert combs. Use two combs if you don’t need to run the fragments out the full length of the gel--it saves time and materials.
5. When the temperature of the agarose solution reaches 60˚C (you should be able to hold the glass for 5 seconds without burning your hand, but make sure the gel hasn't started to polymerise yet), pour the solution into the cast so that the gel is about 2 mm thick. Allow to polymerize (about 45-60 minutes).
6. When polymerized, remove the combs and tape, slide the gel out of the cast, and place in the ethidium bromide bath. A gel must soak for about 30min to be stained.

2% agarose checking gel:
1. Move the gel bath to the bench with the baby gel chamber. Take a gel from the bath with the tongs and place it in the chamber for electrophoreses (make sure the wells are closest to the black electrode). Always make a new gel and put it into the gel bath so that the next person can use it.
2. If necessary, top up the 1x TBE buffer to the fill line of the chamber. The buffer can be reused several times, but should be replaced every second week.
3. Take a microtiter plate and add 1µl of loading dye in a number of wells corresponding to the number of your samples.
4. Add 2.5 µl (if not otherwise stated) of the final PCR product to each well (change tips between samples).
5. Load the first well in the gel with 5 µl of a DNA ladder and 1µl dye, and then the samples (PCR rxn and dye). You can reuse tips here--just place the tip in the buffer in the chamber and pipette up and down to flush the residue.
6. Put the lid on the electrophoresis chamber.
7. Turn on the power supply, adjust the voltage (80 V), and let the gel run for 30-40 minutes.
8. When finished, place the gel on the glass plate in the UV camera box. Be cautious about exposing your skin to UV rays for too long: severe burns may develop without protective clothing or eyewear.
9. When finished, carefully dispose of the gel in the gel waste bin (under the camera). Wipe down the glass with paper towels and dispose of them in the EtBr waste bin.

A general PCR set-up template for keeping notes on the experiment, samples, thermocycler, and master mix.

What are M13 primers, anyways?

A process where we can save money by not having to buy primers that are already dye-labeled, we instead add a 16mer sequence tag (we use the M13F –20 sequence 5'-GTA AAA CGA CGG CCA G-3') onto the 5’ end of one of the primers (called a M13-hybrid primer). After the first couple rounds of PCR cycles, the 16mer tag gets added onto your copied DNA product. We put a small enough amount of the hybrid primer in the PCR cocktail so that it all gets used up by 20-25 cycles. We also put a small amount of M13F-20 primer (16bp in length) in the mix, and this primer has been dye labeled. We then drop the annealing temperature by 5°C and this allows the much shorter dye labeled M13F-20 primer (16bp) to anneal and be added onto the copied DNA strands. We run another 20 cycles and you end up with dye labeled PCR product that is 16bp longer than than your original primer sized product (good to remember if you are comparing it to results using the regular primer).

This is much cheaper because we can buy the M13-20 dye labeled primer in bulk, we need to use so little, and we do not have to buy unique dye labeled primers for every primer set we wish to use (the M13-hybrid primers cost about $10-$12, instead of $70 - $120 for a dye labeled primer).

Sending microsatellite products to the Core facility

3. 9.5μL of HiDi/Liz per well in a 96-well plate.
4. Make a 1/20 dilution of PCR product.

2μL PCR product to 38ul water.

Use a dilution plate.

5. Add 2μL of diluted PCR to the 9.5ul HiDi/Liz in each well.
6. Use a sticky lid and centrifuge.
7. Denature at 95°C for 5min (disable heated lid); DENATURE program.

Place on ice immediately for 5min.

8. Spin down plate and place back on ice.
9. Label plate with name: Wayne_r_<PlateName>, your initials, date.
10. Carry on ice to the Core facility (5th floor Gonda, 5309).
11. Place plate in the bottom shelf of the genotyping refrigerator labeled “Ready to Run with ABI 3700”.
12. Plate results will be on the WebSeq webpage, to be genotyped using GeneMapper.

Microsatellite genotyping using GeneMapper v3.0/3.7

2. Set up marker panel and create a Bin Set in Panel Manager; set up bins if alleles are known. This is done in a hierarchical method in the Panel Manager:

In the left navigation window, click Panel Manager and then “New Kit” (upper far-left button). Name it and select the type of data you want.

Select on the new Kit you created, then click “New Panel”. Name it and press Enter.

Click on the panel you created, then click “New Marker”. Name it, provide marker range, color, and number of repeats, and comments if you choose.

Add in as many markers you have. I create different Kits for each multiplex combination set. Make sure to name everything consistently so you can link up to them later and you know exactly what it means. Example: name Analysis Method, Panel, Kit, etc all matching the microsatellite multiplex primer mix so you can easily cross-reference them when setting up in GeneMapper.

Next, select your Kit and in the Bins menu, select New Bin Set. Name it and then you can select it from the drop-down menu. This Bin Set is used in the Analysis Method to again, be clear about how you name your Kits and Panels.

3. Set up Analysis Method specific to your samples in GeneMapper Manager.

In the GeneMapper Manager, click the Analysis Method Tab, and then click New.

Follow prompts, then name your Method on “General” tab.

“Allele” tab: this is where you link your Analysis Method to the Bin Set you just created. You can also click “Use the marker-specific stutter ratio” if your marker stutters.

If marker is known to stutter, you can the change the stutter ratio (value is a percentage of the time you observe stutter in dataset) and this will avoid “over labeling” of stutter peaks.

“Peak Detector” tab: click “User Specific (rfu)” and here you can set a Minimum Peak Height requirement; useful to avoid over labeling low-intensity peaks. The higher the value, the higher the intensity requirement for peak calling.

The remaining tabs have tons of values you can change to better tweak your analysis. I haven’t figured them all out yet, but don’t let that stop you from having fun!

Click OK, then Done.

4. Select your Analysis Method, Panel, and Size Standard information on Samples Tab of project, and hold “Ctrl-D” to fill down for all samples.

6. On the Genotype Tabs, each individual is represented by a separate entry for each marker it was run with (if you multiplexed markers). Each run is tested for quality control in many categories. The GQ (Genotype Quality) column is most important and is rated one of three quality levels:

Green square: good to go!

Yellow triangle: usually ok to go!

Red hexagon: bad; it’s a no-go!

7. To view all electropheregrams, select all runs in each marker (left navigation window in Genotypes Tab) and click the “Display Plots” button.

8. There are two important buttons in the upper left corner (Peak Selection Mode and Binning Mode buttons). You can view runs and add allele bins here if necessary.

Peak Selection Mode: Here you can click on any peak, add it as an allele you created previous to the analysis run, delete the allele call, or custom name the allele peak as a “new” allele for that individual peak.

Binning Mode: Here you can create/delete/edit bins and increase/decrease the marker range as needed. Any changes made are saved when you exit the screen but you will need to RE-analyze for it to be applied to all the other samples analyzed with that marker. Go back to the Samples Tab and click the “Analyze” button.

9. I recommend viewing all runs to double check the calls made by GeneMapper. If you think a run is good, but the program gave it a GQ red hexagon (failed), you can override this by two ways:

When you make any changes to a run, it automatically replaces quality values with gray triangles. This tells you that you made manual changes to the run.

You can “right” click on the final GQ value and a prompt appears asking “Override the Genotype Quality of this marker?” Yes will also create the gray triangles.

10. Once you have viewed/edited all the runs for a single marker, you can always sort the entries under the Edit menu. You can also edit the Table Settings in Table Settings Editor to hide some of the excessive columns that normally appear on both the Samples and Genotype Tabs. Make sure to select the Table Settings you created in the drop- down menu in order for it to be applied to your project.

11. To Export your Allele Table, click “Export Table” button and save it in your folder. It is saved as a “.txt” file and easily copy/pastes into Excel. No problem!

12. Congratulations! (as a last note, I recommend re-analyzing all samples for a marker when you add more samples for that marker...keeping allele calls consistent and standardized across PCRs).

Troubleshoot:

Check that you are viewing the table in the correct table edit selection. Microsatellite data should be under the Microsatellite Default viewing screen, AFLP with the AFLP screen, and so on.

Check that you have selected the appropriate bin set when viewing alleles. This makes a world of difference.

High resolution melting (HRM) curve analysis protocol

This method is used for quick genotyping of small variants (e.g. indels, SNPs). Primer design is different that of that for microsatellites.
HRM primer design
1. Locate SNP (or other genetic variant) and it’s position using the UCSC Genome Browser or Ensembl in the most recent genome build
2. Obtain about 100 bases of flanking sequence in both directions with the SNP in the middle of the sequence (use the DNA tab to get genomic sequence)
3. Use Primer3 online to submit sequence using default primer picking options

Use the brackets [A] to target the SNP location

4. Design as small of amplicons as possible (target product size of 45-80 bases max)

Setting up the HRM plate and master mix
1. Thaw reagents, found in the pre-PCR room chest freezer

Water

Roche MgCl2

Roche MM (light sensitive)

Primers

2. Make 2μM primer mix (1μL of the forward primer + 1μL of the reverse primer + 98μL of diH2O
3. Make master mix, vortex and spin down
4. Add 2μL of the 20x diluted PCR product and mix by pipetting up and down several times
5. Seal the plate with a sticky lid and spin down using the centrifuge
6. Denature at 95° for 5 min (disable the heated lid) and place on ice immediately (DENATURE programs on most thermocyclers)
7. After 5 min on ice, spin down plate and return to ice
8. Label plate with name using the following format: wayne_r "Plate Name", your initials, date
9. Carry on ice and covered to the Core facility (Gonda 5309)
10. Place denatured plate in genotyping fridge labeled "Ready to Run with ABI 3700" shelf (at the bottom)
11. When the samples have been sequenced, they will be posted on WebSeq and downloaded.

Targeted sequencing

Cloning, colony picking, and PCR

Goal: Use a pipette tip to pick a colony, boil it to lyse plasmid from bacteria, and then PCR it with M13F and R primers to cleanly amplify just the target insert without needed gel band cutout.

Pick & Boil Step
Recommend using a new box of sterile pipette 20 or 200 µl size tips (Caution! Don’t use tips from a box that has been previously opened and used for other work. If you don’t use all the tips, set them aside and label for colony picking use only).

Use a 96-well PCR plate and add 30µl of 0.1X TE buffer into each well you plan to use for a picked colony. We recommend using the bottle of molecular grade 50X TE buffer from which we make the 0.1X solution.

Using a sterile pipette tip, gently touch the selected colony to get some cells on it, then place the tip into the well of the PCR plate. Gently swirl the tip in the well for a few seconds. After you have picked all the desired colonies and “innoculated” the wells of the plate (a tip should be in a well for at least a minute), then gently remove each tip without dripping any liquid into other wells.

Put a flexy silicone lid on the plate and then boil the colonies to spring (lyse) the plasmids from the bacteria. The cycle used should be 95°C for 10 minutes, followed by 4°C forever until you are ready to take the plate out of the PCR machine. Make sure to use the heated lid option!

PCR Reaction Mix & Cycle
You want to put enough plasmid into the PCR reaction to get decent amplification of the insert, but not too much that you have to ultimately cut the insert band out because the plasmid concentration is too high. We have found that 1µl of the boiled colony solution works well to get this result.

Reagents for a 10μL reaction

x1

Qiagen Master Mix

5μL

BSA (10mg/mL; dilute stock using 1μL 100mg/mL stock to 9μL dH2O)

0.4μL

M14F and Reverse primer mix (2μM)

1μL

diH2O

2.6μL

add 9μL of PCR cocktail to 1μL boiled colony per reaction

PCR program
95 ºC for 15 min; 94°C for 30s; (For 45 cycles: 89ºC 30s, 50ºC 60s, 72ºC 60s); 60ºC 30 min; 4ºC forever hold
Do a checking gel to see if you have an insert amplifying in each sample (if there is a ~200bp band, that means there is no insert; if no band, means there was no or very few cells in the boiled solution). Use 3µl of PCR product and 2µl of loading dye for the checking gel. If clean single band of the expected insert size, do the standard next steps for sequencing (Agencourt/Beckman prep, or if UCLA core – Exosap and Big Dye rxn).

1. Remove tubes from incubator, and add 500 µl of buffered phenol. Shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
2. Observe 2 layers. The top layer contains the DNA/RNA and the bottom, organic layer the waste.
3. Remove the top layer and transfer into a new and labeled tube, and discard the bottom layer in the PCI organic waste container.
4. Add 500 µl of PCI to the samples in the new tubes, and shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
5. Observe 2 layers. The top layer contains the DNA/RNA and the bottom, organic layer the waste.
6. Remove the top layer and transfer into a new and labeled tube, and discard the bottom layer in the PCI organic waste container.
7. Add 500 µl of CI to these new tubes, and shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
8. Pipette off the top layer and transfer it to a 1.7ml final epi tube with the final sample name and number.
9. [OPTIONAL] If you want to RNase treat the DNA, add 1ul of a 10 µg/mL stock solution of RNase A to your DNA and incubate at 37ºC for 30-60 min.
10. To this tube add the following:

100 µl of 3M sodium acetate (NaOAc)

1 ml of cold 100% ethanol (EtOH)

11. Invert tubes several times and place in a -20 freezer overnight. (3 hours is sufficient).
12. You can either stop here and do precipitation the next day or after 3 hours, complete the precipitation.

Ethanol Precipitation
1. Remove tubes from the freezer, and centrifuge at max (14,000 rpms for ~10 min) to pellet the DNA.
2. Observe a white or brownish pellet. (May be absent in low concentration samples).
3. Decant the supernatant, being careful not to lose the pellet.
4. Add 1ml of fresh 70% EtOH to the tube containing the pellet. Vortex briefly to re-suspend the pellet.
5. Centrifuge at max (14,000 rpms for 10 min) to pellet the DNA.
6. Decant the EtOH, being careful not to lose the pellet.
7. Remove the remaining 70% EtOH by vacuum centrifuging in the tubes in the Savant Speed-Vac for 10 mins or until the pellet is dry. (Do not use high heat for more than 5 mins).
8. Re-suspend DNA in 100-200 1x TE (or Qiagen’s AE) Buffer.

RNase treatment [if you didn’t do this prior to precipitation]

If you are extracting DNA from an RNAbuffer (e.g. PAXgene RNA tubes which do not contain any DNases), you may consider doing an RNAse treatment of the final DNA*

1. Add 1ul of a 10 µg/mL stock solution of RNase A to your DNA and incubate at 37ºC for 30 min.
2. Recover the DNA by adding 1/10 volume of 3M sodium acetate (pH 6.8) and 2 volumes of isopropanol or 95% ethanol to the DNA containing solution.
3. Incubate on ice for 10 min.
4. Centrifuge at maximum speed for 5 min at room temperature, to pellet the DNA.
5. Discard (carefully) the alcohol. Wash with 70% ethanol and dry DNA via Speed-Vac on low heat for 10 mins.
6. Dissolve in AE, TE or dH2O, as you did in the DNA extraction.

3. Add trial volume of ampure to 20μL of ladder mixture (may need to do more trials if AMPure haven’t been tested in awhile).

Example:

Final concentration of AMPure

Volume of ladder mixture

Volume of AMPure

.2X

20μL

4μL

.4X

20μL

8μL

.6X

20μL

12μL

.8X

20μL

16μL

4. Mix gently with a pipette and incubate at room temperature for 5 minutes.
5. Place tubes in magnetic tube holder and allow to rest for 2 minutes. Beads should be drawn out of solution and should gather on the side of the tube in contact with the magnet.
6. Carefully pipette off supernatant - without disturbing the beads - and discard.
7. Add 500uL fresh 70% ethanol to each tube, then pipette off ethanol and discard.
8. Repeat step 6.
9. Allow beads to air dry with tube caps open to remove all traces of ethanol (~5-10 minutes) or place beads of a 37°C heat block for 3-4 min until dry.
10. Resuspend beads in 10μLuL 1X TE buffer to elute DNA.
11. Transfer elute to new labeled tubes and discard beads.
12. Mix elute with 1μL of loading dye.
13. Electrophorese in 1.5% agarose gel for 50min at 100V.
14. Compare the results of the trial volumes do obtain the most appropriate Xul needed to do the following AMPure Bead Cleanup Protocol.

AMPure XP bead clean-up of DNA

1. Use sample DNA concentration to calculate volume that will yield 1μg of DNA.

To calculate the number of microliters needed to obtain 1μg of DNA, divide 1000 by the concentration of DNA in ng/μL.

To obtain genomic DNA, use a ratio of XuL bead solution per μg of sample DNA (X is the optimum volume figured out by testing the ampure mixture; typically 40-60μL).

2. Obtain clear 1.7mL tubes. In each 1.7mL tube, combine bead solution, DNA, and 1X TE buffer in the following ratio: XμL : 1μg : up to 160μL total volume.

Can be scaled up to combine as much as 3μg DNA with 3XμL bead solution and 1X TE to a total volume of 480μL in each tube.

3. Mix gently with a pipette and incubate at room temperature for 5 minutes.
4. Place tubes in magnetic tube holder and allow to rest for 2 minutes. Beads should be drawn out of solution and should gather on the side of the tube in contact with the magnet.
5. Carefully pipette off supernatant - without disturbing the beads - and discard.
6. Add 500μL fresh 70% ethanol to each tube, then pipette off ethanol and discard.
7. Repeat step 6.
8. Allow beads to air dry with tube caps open to remove all traces of ethanol (~5-10 minutes) or place beads of a 37°C heat block for 3-4 min until dry.
9. Resuspend beads in 10μL 1X TE buffer to elute DNA.
10. Transfer elute to new labeled tubes (combining duplicates from the same initial sample, if possible) and discard beads.
11. NanoDrop to determine new sample concentration. Perform ethanol precipitation to concentrate if necessary.